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Journal of Bacteriology, March 2009, p. 1369-1381, Vol. 191, No. 5
0021-9193/09/$08.00+0     doi:10.1128/JB.01580-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Characterization of a Coxiella burnetii ftsZ Mutant Generated by Himar1 Transposon Mutagenesis ^,

Paul A. Beare,¹ Dale Howe,¹ Diane C. Cockrell,¹ Anders Omsland,¹ Bryan Hansen,² and Robert A. Heinzen^1*

Coxiella Pathogenesis Section, Laboratory of Intracellular Parasites,¹ Electron Microscopy Facility, Research Technologies Branch, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, Hamilton, Montana 59840²

Received 7 November 2008/ Accepted 16 December 2008

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Coxiella burnetii is a gram-negative obligate intracellular bacterium and the causative agent of human Q fever. The lack of methods to genetically manipulate C. burnetii significantly impedes the study of this organism. We describe here the cloning and characterization of a C. burnetii ftsZ mutant generated by mariner-based Himar1 transposon (Tn) mutagenesis. C. burnetii was coelectroporated with a plasmid encoding the Himar1 C9 transposase variant and a plasmid containing a Himar1 transposon encoding chloramphenicol acetyltransferase, mCherry fluorescent protein, and a ColE1 origin of replication. Vero cells were infected with electroporated C. burnetii and transformants scored as organisms replicating in the presence of chloramphenicol and expressing mCherry. Southern blot analysis revealed multiple transpositions in the C. burnetii genome and rescue cloning identified 30 and 5 insertions in coding and noncoding regions, respectively. Using micromanipulation, a C. burnetii clone was isolated containing a Tn insertion within the C terminus of the cell division gene ftsZ. The ftsZ mutant had a significantly lower growth rate than wild-type bacteria and frequently appeared as filamentous forms displaying incomplete cell division septa. The latter phenotype correlated with a deficiency in generating infectious foci on a per-genome basis compared to wild-type organisms. The mutant FtsZ protein was also unable to bind the essential cell division protein FtsA. This is the first description of C. burnetii harboring a defined gene mutation generated by genetic transformation.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The obligate intracellular bacterium Coxiella burnetii causes a zoonotic disease called Q fever that has a near worldwide distribution (22). C. burnetii is highly infectious and is typically transmitted to humans via inhalation of contaminated aerosols associated with domestic livestock operations. Sheep, goats, and dairy cattle are important animal reservoirs (22). Disease in humans normally presents as an acute debilitating influenzalike illness. In rare cases, chronic disease can occur that generally manifests as endocarditis. Acute infections are typically self-limiting and are effectively treated with tetracycline or quinolone antibiotics. Chronic infections are more refractory to antibiotic therapy; however, improved efficacy was recently achieved using a prolonged therapy of doxycycline and hydroxychloroquine (10).

In a eukaryotic host cell, C. burnetii replicates to high numbers in a parasitophorous vacuole having characteristics of a phagolysosome (17, 18). Indeed, the pathogenicity of C. burnetii is associated with its ability to resist the harsh conditions of this intracellular compartment and to survive for extended periods in the extracellular environment (15). Unfortunately, the obligate intracellular nature of C. burnetii imposes considerable experimental constraints in identifying pathogen-associated virulence factors. In fact, lipopolysaccharide is the only currently defined virulence factor of C. burnetii (23). A number of genes encoding potential C. burnetii virulence proteins were revealed upon sequencing the Nine Mile reference strain genome (34). However, lacking a method of gene inactivation, molecular Koch's postulates are impossible to fulfill for these putative virulence factors. As a substitute for genetic manipulation, C. burnetii gene function and regulation has been largely characterized using surrogate hosts, primarily Escherichia coli (16, 24, 40, 42) and more recently Legionella pneumophila (27).

Transformation of C. burnetii was described by Suhan et al. (36) more than 10 years ago. Using a plasmid containing a 5.8-kb C. burnetii autonomous replication sequence, they transformed C. burnetii to ampicillin resistance (4, 35, 36). Transformants exhibited both extrachromosomal replication and integration of the plasmid into the chromosome by homologous recombination. However, a significant problem with the system was the outgrowth of ampicillin-resistant, nontransformed C. burnetii following long-term antibiotic selection. Nonetheless, the study by Suhan et al. showed for the first and only time that foreign DNA could be introduced into C. burnetii by electroporation and that homologous recombination occurs in the organism. Reports of successful transformation showing molecular data have since been published for the obligate intracellular bacteria Anaplasma phagocytophilum (9), Rickettsia monacensis (2), Rickettsia typhi (39), Rickettsia conorii (32), and Rickettsia prowazekii (21, 28-30), with transient expression of recombinant DNA also reported for Chlamydia trachomatis (38). Two of these studies (9, 21) successfully used the mariner family transposon Himar1 to randomly mutagenize the pathogen genome (19). This system relies on transposase-directed random integration of a transposon containing an antibiotic resistance gene for positive selection. Mariner family transposons do not require species specific host factors for efficient transposition and integrate nonspecifically at T/A base pairs (20).

The lack of methods to genetically manipulate C. burnetii significantly impedes progress in understanding the organisms unique intracellular lifestyle and virulence. We show here that the Himar1 transposon system can be used to generate random insertion mutations in the C. burnetii genome. Clonal isolation allowed functional characterization of a C. burnetii transformant harboring a Himar1 transposon insertion in FtsZ, a protein critical for cell division (41). The mutant exhibited septation defects and altered growth kinetics. The mutant FtsZ was also unable to bind FtsA which likely explains the observed phenotypes.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Bacterial strains and plasmids. The bacterial strains and plasmids used in the present study are listed in Table 1. E. coli was grown in Luria-Bertani (LB) broth (33). E. coli transformants were selected on LB agar plates containing either 50 µg of kanamycin (Km)/ml or 10 µg of chloramphenicol (Cm)/ml. C. burnetii Nine Mile phase II (clone 4, RSA439) was propagated in African green monkey kidney (Vero) fibroblasts (CCL-81; American Type Culture Collection) grown in RPMI medium (Invitrogen, Carlsbad, CA) supplemented with 2% fetal bovine serum (FBS) at 37°C in a 5% CO₂ atmosphere. This strain is exempt from U.S. Centers for Disease Control and Prevention select agent regulations (http://www.cdc.gov/od/sap/sap/exclusion.htm) and is suited for work at biosafety level 2 (13).

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TABLE 1. Bacterial strains and plasmids

Construction of Himar1 C9 transposase and transposon plasmids. The restriction enzymes used in the present study were obtained from New England Biolabs (Ipswich, MA). PCR was performed by using Accuprime Pfx or Accuprime Taq (Invitrogen). PCR primers were obtained from Integrated DNA Technologies (San Diego, CA) or Operon Biotechnologies, Inc. (Huntsville, AL), and their sequences are listed in Table 2. All ligations were carried out by using Ligate-it (USB, Cleveland, OH).

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TABLE 2. TaqMan qPCR, PCR and sequencing oligonucleotides

Figure 1A depicts the Himar1 C9 transposase expression plasmid pCR2.1-P1169-Himar1C9, where transposase expression is driven by the C. burnetii CBU1169 promoter (P1169). Himar1C9 from pBVS2-Himar1C9 (generously provided by P. Stewart, Rocky Mountain Laboratories) was amplified by using Accuprime Taq DNA polymerase and the primers Himar1C9-NdeI-F and Himar1C9-HindIII-R and then cloned into pCR2.1-Topo. P1169 was amplified from C. burnetii genomic DNA (gDNA) by using Accuprime Taq and the primers P1169-PstI-F and P1169-NdeI-R and then cloned into the pCR2.1-topo vector, creating pCR2.1-P1169. A PstI-NdeI fragment from pCR2.1-P1169 containing P1169 was subsequently ligated into PstI-NdeI-cut pCR2.1-Himar1C9 to generate pCR2.1-P1169-Himar1C9.

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FIG. 1. Plasmid maps of pCR2.1-P1196-Himar1C9 and p1898-Tn. (A) In pCR2.1-P1196-Himar1C9, the promoter from CBU1169 (P1169) drives expression of the C9 variant of the Himar1 transposase. (B) In p1898-Tn, the Himar1 transposon contains CAT and mCherry genes expressed as a single transcriptional unit from P1169. Also contained within the Himar1 ITRs is a ColE1 origin of replication that allows rescue cloning of the Tn in E. coli. Outside of the transposon is the coding sequence of CBU1898.

Figure 1B depicts the Himar1 transposon plasmid p1898-Tn. BamHI and BspHI restriction sites were incorporated into the pITR-pflgB-GENT (generously provided by P. Stewart) by PCR using the primers pITR-pflgB-GENT-BamHIF and pITR-pflgB-GENT-BspHIR. The subsequent PCR product was ligated together to create pITR-pflgB-GENT-BB. P1169 was amplified from C. burnetii gDNA using P1169-AvaI-F and P1169-NdeI-R, and the resulting PCR fragment was cloned into pCR-BLUNTII-Topo to create pBLUNTII-P1169. A PCR fragment containing the Cm acetyltransferase (CAT) gene was amplified from pEXP1-DEST (Invitrogen) using the primers CAT-NdeI-F and CAT-EcoRV-R and cloned into pCR-BLUNTII-Topo (Invitrogen) to create pBLUNTII-CAT. The NdeI-EcoRV fragment containing CAT was excised and ligated into NdeI-EcoRV cut pBLUNTII-P1169 to create pBLUNTII-P1169-CAT. The AvaI-EcoRV fragment containing P1169-CAT was excised and ligated into AvaI-EcoRV-cut pITR-pflgB-GENT-BB, resulting in the replacement of the flgB promoter and gentamicin gene with P1169-CAT, to produce pITR-P1169-CAT. The mCherry gene was PCR amplified from pcDNA6.2-C-mCherry-DEST (generously provided by S. Grieshaber, University of Florida) using mCherry-BspHI-F and mCherry-BamHI-R primers and cloned into pCR2.1-Topo to create pCR2.1-mCherry. A BamHI fragment containing mCherry was excised and ligated into BamHI cut pITR-P1169-CAT to create pTn. A promoterless version of CBU1898, encoding a C. burnetii IS1111A transposase with 21 copies in the C. burnetii genome, was amplified using CBU1898NotI-F and CBU1898NotI-R and then cloned into pCR-BLUNTII-Topo to create pBLUNTII-1898. A NotI fragment from pBLUNTII-1898 containing CBU1898 was cloned into NotI-cut pTn to create p1898-Tn. Ampicillin, Km, and Cm are not used in the clinical treatment of Q fever (10). Therefore, use of genes that confer resistance to these antibiotics is appropriate for C. burnetii transformation studies, and their use was approved by the Rocky Mountain Laboratories Institutional Biosafety Committee.

Electroporation of C. burnetii and cultivation of transformed bacteria. Large-scale p1898-Tn and pCR2.1-P1169-Himar1C9 plasmid purifications were conducted by using a GenElute HP endotoxin-free plasmid maxiprep kit (Sigma-Aldrich, St. Louis, MO) and concentrated by using a Montage PCR filter unit (Fisher Scientific, Pittsburgh, PA). C. burnetii was purified from infected cells by renografin density gradient centrifugation as previously described (5), washed twice in Coxiella transformation buffer (CTM; 272 mM sucrose, 10% glycerol), and then resuspended in CTM at approximately 2.5 x 10¹¹ genome equivalents (GE) per ml. C. burnetii GE were enumerated via TaqMan quantitative PCR (qPCR) as previously described (6). To 50 µl of a C. burnetii suspension, 3 µl (12 µg) of p1898-Tn and 4 µl (14 µg) of pCR2.1-P1169-Himar1C9 were added. The cell suspension was placed in a 0.1-cm gap electroporation cuvette on ice and electroporated with a field strength of 16 or 20 kV/cm, a resistance of 500 , and a capacitance of 25 µF using an ECM630 Electro-Cell manipulator (BTX, Holliston, MA). As controls, C. burnetii was electroporated without DNA or mock electroporated with DNA. For first-round electroporations, 950 µl of RPMI was added directly to the cuvette, and 100 µl of the mixture was used to infect confluent Vero cells in one well of a six-well plate. For the second-round electroporations, the entire 1 ml was used to infect confluent Vero cells in a T-75 flask. Infected Vero cell cultures were incubated for 2 h at room temperature with gentle rocking, and then RPMI supplemented with 2% FBS was added to cell cultures, which were incubated for an additional 22 h. Cm was then added to the media at a final concentration of 5 µg/ml. After 1 week, the cell monolayers were harvested by scraping and disrupted by sonication. The sonicate was centrifuged at 1,000 x g for 5 min to pellet host nuclei and large cell debris, and the supernatants were used to infect new Vero cell monolayers. This process was repeated every 1 to 2 weeks. During all cell culture, the medium was removed every 3 to 4 days and replaced with fresh medium containing antibiotic.

Isolation and whole-genome amplification of gDNA. C. burnetii gDNA was isolated by using either a PowerMicrobial Midi or a Ultraclean Microbial DNA isolation kit (Mo Bio, Carlsbad, CA) with an additional heating step (85°C for 30 min) prior to the physical disruption of the cells. Plasmid DNA was isolated by using a Qiaprep Spin miniprep kit (Qiagen, Valencia, CA) or a GenElute HP endotoxin-free plasmid maxiprep kit. gDNA was whole-genome amplified using 8 µl of purified gDNA as a template and the Illustra GenomiPhi V2 DNA amplification kit (GE Healthcare, Piscataway, NJ).

PCR and Southern blot detection of integrated transposons. Detection of the Tn was conducted by amplifying CAT, mCherry, and CAT-mCherry genes using the primer pairs CAT-NdeI-F/CAT-EcoRV-R, mCherry-BspHI-F/mCherry-BamHI-R and CAT-NdeI-F/mCherry-BamHI-R (Table 2), respectively. For Southern blots, gDNA or whole-genome amplified DNA was digested with BamHI and BsaHI or with BsaHI alone and then separated on a 0.8% agarose gel by electrophoresis. Digested gDNA was transferred by blotting to Hybond N+ membranes (GE Healthcare) as described by Sambrook et al. (33), except that the transfer medium used was 0.4 M NaOH. Probe DNA specific to CAT or mCherry genes was generated by PCR using the primer pairs CAT-NdeI-F/CAT-EcoRV-R and mCherry-BspHI-F/mCherry-BamHI-R, respectively. Probe DNA specific to ftsZ (CBU0141) was generated using the primer pair CBU0141-F/CBU0141-R. A probe specific to the 1-kb Plus DNA marker (Invitrogen) was also used. Probe DNA (200 ng) labeling and subsequent blot hybridizations were conducted according to the instructions and using the reagents provided for a Gene Images AlkPhos direct labeling and detection kit (GE Healthcare).

Rescue cloning and sequencing of Tn integration sites. gDNA was isolated from transformed C. burnetii and digested with ClaI or PsiI for 4 h at 37°C. Digested gDNA was heated at 65°C for 20 min and ligated together. E. coli Top10 cells (Invitrogen) were transformed with ligated DNA and plated on LB agar containing 10 µg of Cm/ml. Colonies containing rescued plasmid with a ColE1 origin of replication were grown overnight in LB broth containing 10 µg of Cm/ml. Plasmid DNA was isolated and sequenced using the sequencing primers ColE1-3'-out and Cm-5'-out (Table 2) to obtain the genomic Tn integration site. Sequences were analyzed by using VectorNti 10 advance (Invitrogen) and the BLAST algorithm (http://blast.ncbi.nlm.nih.gov/Blast.cgi).

Clonal isolation of a C. burnetii Tn mutant. Vero cells cultivated on 12-mm glass coverslips in a 24-well tissue culture plate were infected with Tn mutagenized C. burnetii. At 5 days postinfection, coverslips were transferred to a 90-mm-diameter petri dish containing approximately 10 ml of RPMI medium, and individual parasitophorous vacuoles were extracted by micromanipulation as previously described (3). The harvested content from individual vacuoles was mixed with 200 µl of RPMI supplemented with 2% FBS, and the suspension was used to infect Vero cells in one well of a six-well tissue culture plate. C. burnetii transformant clones were subsequently expanded in Vero cells in the presence of Cm (5 µg/ml) and purified as described above.

Reverse transcription-PCR (RT-PCR) detection of CAT and mCherry transcripts. Confluent Vero monolayers in six-well plates were infected with C. burnetii and cells lysed for RNA extraction at 3 and 7 days postinfection. Infected cells were washed twice with phosphate buffered saline (PBS; 1 mM KH₂PO₄, 155 mM NaCl, 3 mM Na₂HPO₄ [pH 7.4]) and lysed with 1 ml of TRIzol (Invitrogen), and then lysates were homogenized twice for 40 s at setting 5 in a FastPrep FP120A instrument using FastRNA Pro Blue vials (MP Biomedicals, Irvine, CA). To each sample, 200 µl of 1-bromo-3-chloropropane (Sigma-Aldrich) was added, and the mixture vortexed for 15 s, followed by heating for 10 min at 65°C. Lysates were subsequently centrifuged for 15 min at 12,000 x g at 4°C, the aqueous phase was removed, and RNA was extracted by using an RNeasy kit (Qiagen). The RNA yield was determined by spectrophotometry (A₂₆₀/A₂₈₀), and the integrity was verified by using Agilent 2100 Bioanalyzer (Agilent Technologies, Santa Clara, CA). A total of 50 ng of each RNA was treated with DNase I, reverse transcribed into cDNA, and amplified by using a CellsDirect One-Step qRT-PCR kit (Invitrogen) according to the manufacturer's protocol. A minus reverse transcriptase control was conducted by using Platinum Taq DNA polymerase (Invitrogen) in place of the Superscript III-Platinum Taq mix in a CellsDirect One-Step qRT-PCR kit. The expression of the CAT and mCherry genes was quantified by qPCR using primers and probes (Table 2) specific for these genes.

Light and immunofluorescence microscopy. For immunofluorescence labeling, Vero cells on 12-mm glass coverslips in a 24-well tissue culture plate were infected with C. burnetii for 5 days followed by fixation in 100% methanol for 5 min. The C. burnetii parasitophorous vacuole membrane was stained using a monoclonal antibody directed against human CD63 (BD Pharmingen, San Jose, CA) and an Alexa Fluor 488 goat anti-mouse immunoglobulin G (IgG) (Invitrogen) antibody. For live-cell imaging, Vero cells on glass-bottom 35-mm petri dishes were infected for 5 days and then viewed by differential interference contrast (DIC) and confocal fluorescence microscopy. Microscopy was conducted with a modified Perkin-Elmer UltraView spinning disc confocal system connected to a Nikon Eclipse Ti microscope equipped with a Photometrics Cascade:512F digital camera (Roper Scientific, Tucson, AZ). Confocal fluorescent (0.2-µm sections) and DIC images were acquired by using Metamorph software (Universal Imaging, Dowingtown, PA). All images were processed by using ImageJ software (written by W. S. Rasband at the U.S. National Institutes of Health, Bethesda, MD [http://rsb.info.nih.gov/ij/]) and Adobe Photoshop (Adobe Systems, San Jose, CA).

Quantification of C. burnetii infectivity and replication. C. burnetii infectivity for Vero cells was quantified by using a fluorescent focus-forming unit assay as previously described (6). The kinetics of C. burnetii replication in Vero cells were quantified as previously described (6).

SEM. Scanning electron microscopy (SEM) was performed on C. burnetii that were purified from Vero cell monolayers at 7 days postinfection. Bacteria were fixed on silica chips (Ted Pella, Inc., Redding, CA) overnight at 4°C with 2.5% glutaraldehyde, 4% paraformaldehyde, and 0.05% sucrose in a 0.1 M sodium cacodylate buffer (pH 6.8). Cells were then postfixed in 0.5% reduced osmium using a Pelco Biowave microwave (Ted Pella, Inc.) at 250 W 2X (2 min on, 2 min off, 2 min on) under 15 in Hg vacuum. The samples were ethanol dehydrated for 45 s in a microwave under vacuum at 250 W and critical point dried in a Bal-Tec cpd 030 drier (Balzers, Pell City, AL). Chips were then coated with 75 Å of iridium in an IBS ion beam sputterer (South Bay Technology, Inc., San Clemente, CA) and imaged by using a Hitachi S-5200 In-Lens SEM (Hitachi, Pleasanton, CA).

Expression of recombinant FtsZ, FtsZ::Tn, and FtsA. Full-length C. burnetii ftsZ and ftsA were amplified by PCR using the primer pairs CBU0141-F/CBU0141-R and CBU0140-F/CBU0140-R, respectively. ftsZ::Tn was amplified using the primer pair CBU0141-F/CBU0141-B2-R (Table 2). PCR products were cloned into pENTR-D-Topo (Invitrogen), and the ftsZ and ftsZ::Tn plasmid clones were subsequently used in a LR clonase II reaction with pEXP1-DEST (Invitrogen) to generate plasmids encoding N-terminally Xpress-tagged protein (Table 1). The ftsA plasmid clone was used in an LR clonase II reaction with pDEST-15 (Invitrogen) to generate a plasmid encoding an N-terminally glutathione S-transferase (GST)-tagged protein (Table 1). The pEXP1-ftsZ, pEXP1-ftsZ::Tn, and pDEST-15-ftsA plasmids were used as templates in in vitro transcription/translation (IVTT) reactions using the RTS 100 E. coli HY kit (Roche, Indianapolis, IN) to achieve expression of cell-free recombinant protein. Total protein from IVTT reactions was separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) on a 10% gel and transferred to a nitrocellulose membrane (Bio-Rad, Hercules, CA). After transfer, membranes were blocked for 1 h at room temperature in Tris-buffered saline (TBS; 150 mM NaCl, 100 mM Tris-HCl [pH 7.6]) containing 0.1% Tween 20 and 5% nonfat milk. Membranes were then incubated for 1 h at room temperature in TBS-Tween 20 containing a mouse monoclonal anti-Xpress (Invitrogen) or a goat polyclonal anti-GST antibody (GE Healthcare). Membranes were washed and incubated for 1 h at room temperature in TBS-Tween 20 containing anti-mouse or anti-goat immunoglobulin G secondary antibody conjugated to horseradish peroxidase (Pierce, Rockford, IL). Reacting proteins were detected via enhanced chemiluminescence using SuperSignal West Pico reagent (Pierce).

FtsZ pull-down assays. An IVTT reaction containing GST-FtsA was mixed with an IVTT reaction containing equal amounts of either Xpress-FtsZ or Xpress-FtsZ::Tn, and the mixtures were incubated for 2 h at 37°C. Glutathione-Sepharose 4B (GE Healthcare) was added to IVTT reaction mixtures, which were then incubated at room temperature for 1 h. Glutathione-Sepharose was pelleted by centrifugation at 500 x g for 5 min and washed five times in TBS containing 1% Triton X-100. The glutathione-Sepharose was resuspended in SDS-PAGE loading buffer and boiled for 10 min. Proteins were separated by SDS-PAGE on a 10% gel and transferred to a nitrocellulose membrane (Bio-Rad). Immunoblotting was conducted as described above using anti-Xpress to detect the presence of Xpress-FtsZ or Xpress-FtsZ::Tn.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Construction of a C. burnetii-optimized Himar1 transposon system. Chromosomal transposition of the Himar1 transposon occurs in E. coli when the transposon and unregulated Himar1 transposase are placed on the same plasmid. This results in low yields of the original construct (P. Stewart, unpublished data). Therefore, we placed these elements on separate plasmids and used a two-plasmid approach to transform C. burnetii. The promoter for the CBU1169 gene (P1169), which encodes the small heat shock protein Hsp20, was chosen to drive expression of both the Himar1 C9 transposase variant and the CAT and mCherry genes within the Himar1 transposon. Previous microarray and qPCR studies in our lab showed that the CBU1169 promoter is active throughout the C. burnetii infectious cycle (data not shown). The Himar1 C9 transposase plasmid pCR2.1-P1169-Himar1C9 contains the CBU1169 promoter immediately upstream of the Himar1 C9 transposase gene (Fig. 1A). The Himar1 transposon plasmid p1898-Tn contains a ColE1 origin of replication, CAT and mCherry red fluorescent protein-coding genes flanked by the transposon ITR elements (Fig. 1B). The CAT and mCherry genes both have the CBU1169 RBS directly upstream of their respective start codons, which allows expression of both genes as a single transcriptional unit from P1169 upstream from the CAT gene. Resistance to Cm was chosen as a method of positive selection since the antibiotic is efficacious in cell culture models of C. burnetii infection (31) and is not used in the clinical treatment of Q fever (10). mCherry was chosen as a fluorescent marker instead of green fluorescent protein because C. burnetii autofluoresces green when excited with light in the 488-nm range (unpublished data). A promoterless copy of CBU1898, encoding an IS1111A transposase with 21 copies in the C. burnetii genome, was inserted into a NotI site outside of the Himar1 transposon to promote homologous recombination of the transposon plasmid into the C. burnetii chromosome in the event that transposition was unsuccessful.

Himar1 transposon mutagenesis. Prior to initiating electroporation experiments, the concentration of Cm required to inhibit C. burnetii growth in Vero cells was established. Vero cells were infected with C. burnetii in the presence of 0.5, 2, 5, or 10 µg of Cm/ml. At 5 days postinfection, focus-forming units were not detected in cell cultures treated with 5 or 10 µg of Cm/ml (data not shown). Thus, 5 µg/ml was chosen as the concentration of Cm for the selection of transformants. C. burnetii was electroporated with p1898-Tn and pCR2.1-P1169-Himar1C9 using a field strength of 16 kV/cm as described in Materials and Methods. Ten percent of the electroporated C. burnetii, termed B2, was used to infect one well of a six-well plate containing approximately 10⁶ Vero cells. Controls included C. burnetii exposed to the same amount of both plasmids, but not electroporated, and C. burnetii electroporated in the absence of DNA. At 24 h postinfection Cm was added to the growth media for selection. Every 1 to 2 weeks infected Vero cells were harvested, disrupted gently by sonication, and released C. burnetii used to infect new monolayers under constant Cm selection. After five C. burnetii passages in Vero cells, organism-containing parasitophorous vacuoles were clearly visible by phase-contrast light microscopy only in cells infected with organisms electroporated in the presence of both plasmids. gDNA was isolated from cell cultures and PCR conducted to detect the presence of full-length genes encoding CAT (606 bp) and mCherry (708 bp), and the CAT-mCherry region (1,472 bp) of p1898-Tn. Predicted PCR products were observed only in DNA extracted from cultures infected with B2 (data not shown). To determine whether the p1898-Tn DNA of B2 transformants had integrated by homologous recombination via CBU1898 or was autonomously replicating, PCR of the region from CBU1898 to the CAT gene (using primers CBU1898-NotI-F and CAT-EcoRV-R; Table 2) was conducted. A predicted PCR product of 2.3 kb was not detected (data not shown), indicating that transposition of the Himar1 transposon had occurred.

B2 transformants were then expanded, and gDNA was isolated from purified bacteria. Rescue cloning of the Tn ColE1 origin of replication was conducted by cutting B2 gDNA with ClaI, an enzyme that does not cut the Himar1 Tn but has 1,121 cut sites throughout the C. burnetii genome. ClaI-digested B2 gDNA was self-ligated and used to transform E. coli. Plasmid DNA was isolated from 36 Cm-resistant colonies and sequenced to establish the Tn integration site. All clones were identical, with a Tn integration in the 3' end of CBU0141 that encodes the cell division protein FtsZ (Fig. 2A).

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FIG. 2. Analysis of the ftsZ Tn insertion of C. burnetii B2c. (A) Schematic of the C. burnetii ftsZ chromosomal region in wild-type C. burnetii (NMII) bacteria and the B2c clone. The Tn is flanked by ITR elements (black arrowheads). The binding sites for PCR primers 1 (CBU0141-F), 2 (P1169-NdeI-R), and 3 (CBU0141-R) are shown. Chromosomal regions of NMII and B2c corresponding to the ftsZ probe are depicted. The locations of BsaHI restriction sites and the predicted restriction fragment sizes resulting from a chromosomal BsaHI digest are indicated below the NMII and B2c chromosomal maps. (B) PCR primers 1 and 3 amplified wild-type ftsZ (1,179 bp) from gDNA of NMII and B2, but not B2c. A 3,698-bp product indicating Tn insertion into ftsZ was amplified from B2 and B2c gDNA, but not NMII gDNA. PCR primers 2 and 3 amplified a 594-bp product from gDNA of B2 and B2c, but not NMII, confirming Tn insertion into ftsZ of B2 and B2c and also revealing the Tn orientation (A) in the chromosome. (C) Southern blot of BsaHI-digested gDNA from NMII, B2 and B2c probed for ftsZ. The ftsZ probe hybridized with a 5,540-bp band in gDNA from B2 and B2c gDNA, but not NMII. This fragment corresponds to the 2,734-bp fragment hybridized in NMII gDNA, shifted by 2,806 bp due to the insertion of the Tn into ftsZ.

The identification of 36 identical ftsZ transposon insertions by rescue cloning did not necessarily preclude the existence of other Tn insertions within B2 C. burnetii. Therefore, we derived a clonal population of C. burnetii, termed B2c, from B2 using micromanipulation (3). PCR of B2 gDNA using ftsZ-specific primers resulted in only two bands: an 3.7-kb band representing ftsZ with the integrated Tn and an 1.2-kb band representing full-length ftsZ (Fig. 2B). Tn-disrupted ftsZ was also detected in B2c gDNA, but full-length ftsZ was not. These data indicated B2c was clonal for ftsZ::Tn and that C. burnetii transformants harboring other Tn insertions were absent. The presence of full-length ftsZ in the B2 sample indicated that organisms containing additional Tn insertions were present and/or that B2 cell cultures still contained carry over nontransformed C. burnetii. Transposition of the Tn in ftsZ of B2 and B2c was also confirmed by Southern blotting (Fig. 2C).

Optimization of Himar1 transposon mutagenesis. We identified a single Himar1 transposon insertion in our initial experiment. Therefore, we tested a higher electroporation field strength (20 kV/cm) to determine whether transformation efficiencies could be improved. In addition, we increased the number of Vero cells used in the initial C. burnetii infection to 8 x 10⁶ cells and, in most cases, infected cells with the entire electroporated sample. Seven electroporation experiments were conducted with C. burnetii electroporation experiments 1, 2, 3, 6, and 7 using a field strength of 16 kV/cm and electroporation experiments 4 and 5 using a field strength of 20 kV/cm. T-75 flasks containing Vero cell monolayers were infected with all electroporated C. burnetii with the exception of electroporation experiment 7, where two T-75 flasks were each infected with one-half of electroporated organisms (experiments 7a and 7b). At 24 h postinfection, 5 µg of Cm/ml was added to each flask for selection. Infected Vero cell monolayers were disrupted, and new monolayers were infected every 1 to 2 weeks under constant antibiotic selection. Consistent with the previous experiment, parasitophorous vacuoles containing Cm-resistant C. burnetii were clearly visible following the fifth passage of C. burnetii transformants in Vero cells. All flasks were positive by PCR for CAT, mCherry, and CAT-mCherry regions (data not shown). Total gDNA (host and bacterial) was extracted from infected cells corresponding to each electroporation experiment and whole genome amplified. Amplified DNA was digested with ClaI or PsiI, enzymes that do not cut within the transposon, and the digested DNA was self-ligated to allow rescue cloning of the Tn ColE1 origin of replication. A total of 94 Cm-resistant colonies were obtained, with 55 and 39 colonies recovered from PsiI- and ClaI-digested DNA, respectively. Table 3 lists the 34 unique Tn integration sites revealed by sequencing rescue-cloned plasmid DNA from each electroporation experiment. The sizes of the plasmids ranged from 3,416 bp to more than 13,000 bp. Twenty-nine Tn insertions were within coding regions, while five Tn insertions were intergenic. Two insertions were found in the QpH1 plasmid. Disrupted coding regions included 16 encoding proteins with predicted functions and 13 encoding hypothetical proteins. Insertions into seven genomic sites (CBU1745, CBU2021, CBU0921, CBU1430, CBU1701, and two intergenic regions) were common to both ClaI- and PsiI-derived rescue clones, indicating restriction sites of both enzymes reside close to the Tn integration site (Table 3). No common Tn insertion events were observed between the seven different electroporation experiments. Electroporation experiments 4 and 5, where organisms were electroporated at the higher field strength (20 kV/cm), yielded the highest number of rescue clones (6 and 13, respectively).

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TABLE 3. Location of Tn insertions in C. burnetii transformants

Figure 3 shows a Southern blot of amplified gDNA isolated from C. burnetii associated with each electroporation experiment. DNA was digested with BsaHI, an enzyme that does not cut within the Tn, and BamHI, which has two cut sites within the Tn generating a single 775-bp band containing the mCherry gene. The 775-bp band was evident in all lanes containing DNA from electroporated C. burnetii. Bands greater than 1.2 kb were also observed and corresponded to BamHI/BamHI or BamHI/BsaHI fragments containing the ITR-p1169-CAT region of p1898-Tn (1.2 kb) plus flanking DNA. Based on the C. burnetii Nine Mile genome sequence, the expected sizes for the BamHI/BamHI or BamHI/BsaHI fragments detected by the CAT probe for each of the 34 mapped Tn insertions were established (see Table S1 in the supplemental material). In general, a band on the Southern blot could be associated with the expected size of the disrupted restriction fragment that resulted from each identified insertion event. For example, in electroporation experiment 1, Tn insertions disrupted CBU0573, CBU0964, CBU1196, CBU1745, and CBU2020 and bands of the expected sizes (1,614, 1,768, 1,729, 2,961, and 1,631 bp, respectively) were evident on the Southern blot (Fig. 3C). The Southern blot also revealed band sizes that could not be associated with a Tn insertion identified by rescue cloning. These bands likely correspond to Tn insertions too far from a ClaI or PsiI restriction site to be efficiently rescue cloned or represent fragments that are toxic in E. coli.

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FIG. 3. Southern blot analysis of C. burnetii transformants. (A) A schematic of C. burnetii gDNA containing a Tn integration. The regions detected by the CAT and mCherry probes are demarked by solid black bars. The location of cut sites for BamHI and BsaHI and the predicted sizes of bands detected by the CAT (>1,223 bp) or mCherry (775 bp) probes are shown. The dotted vertical line indicates the border of the Tn and shows the smallest possible BamHI-BamHI or BamHI-BsaHI restriction fragment size (1,223 bp) that could be detected by the CAT probe. Double parallel lines indicate that the distance between the BamHI site within the Tn and the flanking BamHI or BsaHI site will vary depending on the insertion site of the Tn. (B) Southern blot profiles of untransformed C. burnetii (NMII) gDNA and whole genome amplified DNA from electroporation experiments 1, 2, 3, 4, 5, 6, 7a, and 7b digested with BamHI and BsaHI and probed for CAT and mCherry genes. (C) Southern blot profile of electroporation experiment 1 showing fragments detected by the CAT probe whose sizes correspond to Tn insertions within CBU1745, CBU0964, CBU1196, CBU2020, and CBU0573 recovered by rescue cloning. The 775-bp fragment detected by the mCherry probe is also indicated. DNA molecular weight marker (M) sizes are indicated.

Characterization of the B2c ftsZ::Tn mutant. The clonal isolation of the B2c transformant allowed analysis of CAT and mCherry gene expression during the C. burnetii infectious cycle and characterization of the mutant's phenotype. Vero cells were infected with B2c and RNA extracted at 3 and 7 days postinfection. By quantitative RT-PCR, CAT gene expression of B2c was determined to be 3-fold higher than mCherry gene expression at each time point (data not shown). The lower expression of the mCherry gene may be due to its downstream location relative to the CAT gene on the transcriptional unit contained on the Tn. Nonetheless, transcription of the mCherry gene correlated with red fluorescent organisms in live (Fig. 4A) and fixed (Fig. 4B) Vero cells at 5 days postinfection.

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FIG. 4. Microscopy of B2c-infected Vero cells. (A) Vero cells were infected with B2c for 5 days and then imaged live by DIC and confocal fluorescence microscopy. A confocal Z series of B2c-infected cells was taken at 568 nm, merged, then overlaid onto the DIC image. Red fluorescent C. burnetii are apparent in infected cells. (B) Confocal fluorescence image of a Vero cell infected with B2c for 5 days, fixed with methanol, and then immunostained for the lysosomal protein CD63 (green). Bars, 5 µm.

FtsZ is highly conserved in eubacteria and plays an essential regulatory and structural role in bacterial cell division (41). Therefore, we examined B2c for potential growth defects. Growth of B2c and wild-type C. burnetii in Vero cells was quantified by qPCR over 7 days in the absence of Cm. B2c had a generation time during exponential phase of 19.8 h with a net 1.63-log increase in GE over 7 days. This contrasted with a generation time of 11.7 h and a net 2.51-log increase in GE for wild-type organisms (Fig. 5). B2c growth kinetics were similar in cultures containing 5 µg of Cm/ml (data not shown). The slower growth of B2c relative to wild-type organisms correlated with a higher percentage of B2c organisms containing a division septa (45.6% versus 8.2%, respectively) (Fig. 6). Six percent of B2c cells also contained more than one division septae, a morphology not seen with wild-type organisms. The occurrence of filamentous forms of B2c with incomplete septae and presumably containing more than one genome was associated with a lower infectivity of B2c for Vero cells than control organisms on a per genome basis. Approximately 2.5 more B2c than wild-type C. burnetii GE were required to generate a single infectious foci in Vero cells (data not shown). Collectively, these data indicate that the Tn insertion in B2c disrupts FtsZ function and consequently cell division.

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FIG. 5. Growth kinetics of B2c. DNA was extracted from Vero cells infected with B2c or wild-type C. burnetii (NMII) at 0, 1, 2, 3, 5, and 7 days postinfection. B2c had a slower generation time than NMII (19.8 h versus 11.7 h) during exponential phase (2 to 5 days postinfection). This correlated with a 1.63-log increase in GE over 7 days compared to a 2.51-log increase for NMII. GE between B2c and NMII were significantly different (P < 0.005) from 2 to 7 days postinfection according to the Student t test.

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FIG. 6. Morphological analysis of B2c. B2c and wild-type C. burnetii purified from Vero cells at 7 days postinfection were visualized by SEM. (A) Low (left) and high (right) magnifications of wild-type organisms (top) and B2c (bottom). (B) Histogram showing the percentage of B2c and wild-type C. burnetii with no division septum, one division septum, or more that one division septum (n > 250). Representative SEM images of the three scored morphologies are shown, with septae indicated by white arrowheads. B2c cells exhibited more division septae than wild-type organisms.

Binding properties of B2c FtsZ::Tn. The Tn insertion in the ftsZ gene of B2c results in substitution of the FtsZ C-terminal 25 amino acids (aa) with 17 Tn-encoded amino acids (Fig. 7). The C terminus of native FtsZ contains a motif consisting of 8 aa [L(D/E)(I/V)PX(F/Y)(L/I)(R/K)] that is essential for binding of the membrane-associated protein FtsA and localization of FtsZ polymers to the plasma membrane (41). Therefore, we examined the ability of wild-type C. burnetii FtsZ and B2c FtsZ containing the 17 Tn-encoded amino acids to bind FtsA. Fusion proteins corresponding to FtsZ, FtsZ::Tn and FtsA with predicted molecular masses of 47, 45, and 73 kDa, respectively, were produced by IVTT (Fig. 8A). IVTT lysates were mixed together, and GST pull-down assays were performed. As shown by immunoblotting, GST-FtsA only bound wild-type FtsZ (Fig. 8B). Thus, the growth defects of B2c are associated with the inability of FtsZ::Tn to bind FtsA.

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FIG. 7. Domain structure of E. coli FtsZ and alignment with wild-type C. burnetii (NMII) and B2c FtsZ. (A) Diagram of the domain structure of E. coli FtsZ. The N-terminal domain (black) is involved in the formation of the Z ring which is essential for cell division. The globular C-terminal domain contains a conserved motif (gray) that is essential for binding of FtsA and ZipA. The location of the Tn insertion in B2c FtsZ is marked by a triangle. (B) CLUSTAL W alignment of E. coli FtsZ, NMII FtsZ and B2c FtsZ. The FtsA binding region of E. coli FtsZ is highlighted in gray. Tn-encoded amino acids of the B2c FtsZ protein are italicized.

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FIG. 8. FtsZ pull-down assay. (A) Production of C. burnetii Xpress-FtsZ, Xpress-FtsZ::Tn, and GST-FtsA in IVTT reactions. An aliquot of each IVTT reaction was separated by SDS-PAGE, and fusion proteins were detected by immunoblotting with anti-Xpress or anti-GST antibodies. Bands of approximately 47 and 45 kDa were detected by the anti-Xpress antibody and correspond to Xpress-tagged FtsZ and FtsZ::Tn, respectively. An 73-kDa band was detected with anti-GST antibody and corresponds to GST-tagged FtsA. Molecular mass marker sizes are shown. (B) Pull-down assay using GST-FtsA. An IVTT reaction containing either Xpress-FtsZ or Xpress-FtsZ::Tn was mixed with an IVTT reaction containing GST-FtsA, and then the pull-down procedure was conducted with GST-Sepharose. Pull-downs of Xpress-FtsZ, Xpress-FtsZ::Tn, or GST-FtsA IVTT reactions alone were conducted as controls. GST-FtsA bound to only wild-type C. burnetii FtsZ, as revealed by immunoblotting for the Xpress epitope tag.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The study of C. burnetii has been significantly hampered by the inability to genetically manipulate the organism. We describe here the use of a mariner-based Himar1 transposon for generating random mutations within the C. burnetii genome. Successful transposon mutagenesis allowed the cloning and characterization of a C. burnetii ftsZ::Tn mutant, the first such gene-specific mutant of this organism derived by genetic transformation. Given the slow growth rate of the B2c FtsZ::Tn mutant relative to wild-type C. burnetii, the isolation of this mutant in our first-round electroporation experiment was unexpected. Indeed, all 36 B2-derived rescue clones contained the same Tn insertion, although the existence of other Tn insertions in the population could not be excluded.

The B2c clone contains a Himar1 transposon insertion that results in FtsZ lacking its C-terminal 25 aa. FtsZ is a bacterial tubulin-like protein that has two functional domains. The highly conserved N-terminal domain (300 aa) is associated with FtsZ polymerization, a process required for formation of a cytokinetic septal ring (termed the Z ring). The well-conserved C-terminal domain (15 aa) binds the membrane-associated cell division proteins FtsA and ZipA. These proteins, in turn, recruit downstream cell division proteins (41). As shown in pull-down experiments, B2c FtsZ::Tn does not bind FtsA, a binding deficiency that likely extends to C. burnetii ZipA (CBU0541). Thus, B2c FtsZ::Tn is presumably deficient in cytoplasmic membrane targeting and consequently cell division, a hypothesis consistent with the mutant's slow generation time relative to wild-type C. burnetii and appearance as filamentous forms containing multiple division septae. An E. coli FtsZ-null mutant is not viable, while an FtsZ C-terminal truncation mutant retains the ability to form a Z ring (41). B2c organisms still divide; thus, their truncated FtsZ must function in the absence of FtsA/ZipA binding. Another possibility is that B2c FtsZ::Tn uses a different protein to anchor to the membrane. Indeed, some organisms produce FtsZ with the conserved C-terminal domain but lack obvious FtsA or ZipA homologues (8).

In our optimized second-round electroporation experiments, 34 additional Tn insertions were identified by rescue cloning. Thirty-two are spread throughout the C. burnetii chromosome, while two reside within the large (37.4 kb) QpH1 plasmid. Based on Southern blot results, other Tn insertions are clearly evident that were not rescue cloned. Lack of cloning may result from large ClaI or PsiI restriction fragments that are inefficiently cloned or clones that are toxic to E. coli. Tn insertions were found in 16 genes encoding proteins with predicted function. Insertions predicted to disrupt the function of six genes are of note due to their potential deleterious effects. CBU0003 encodes RecF, a multifunctional protein with roles in recombinational DNA repair, homologous genetic recombination, and DNA replication (7). CBU0036 encodes a protein with homology to FabZ, a β-hydroxymyristoyl-ACP dehydratase that is involved in fatty acid biosynthesis. C. burnetii encodes two FabZ-like proteins (CBU0036 and CBU0614) that may be functionally redundant. CBU0573 encodes a medium/long-chain specific acyl-coenzyme A (CoA) dehydrogenase that is involved in the breakdown of CoA-conjugated fatty acids that stem from β-oxidation and/or amino acid metabolism (11). CBU1196 encodes ClpA, an ATP-dependent Clp protease that binds ClpP protease. The subsequent ClpAP complexes are involved in protein degradation and disaggregation (43). CBU1430 encodes tRNA pseudouridine synthase B (TruB) that is involved in production of pseudouridine. Pseudouridine residues are present in most RNAs (25) and are commonly found in functionally important regions of RNA (1, 12). CBU1410 encodes citrate synthase, the first and rate-limiting enzyme of the tricarboxylic acid (TCA) cycle. C. burnetii preferentially transports and metabolizes TCA cycle intermediates and precursors in axenic media (14, 26), suggesting that the TCA cycle is important for C. burnetii energy production. Interestingly, Agrobacterium tumefaciens citrate synthase mutants have attenuated virulence, reduced type IV secretion gene expression, and slightly reduced growth in minimal media (37).

The Tn insertions identified in structural genes obviously do not disrupt protein functions required for C. burnetii growth in Vero cells. The five Tn insertions identified in intergenic regions, while not directly disrupting protein coding, may still affect expression of flanking genes. For example, the Tn insertion 4 bp upstream of CBU0114, encoding the protein YajQ, may affect expression of the gene due to disruption of its ribosome-binding site and promoter.

An electroporation field strength of 20 kV/cm yielded a larger number of unique second-round transposon insertions than electroporations conducted at 16 kV/cm. The transformation frequency of our electroporation procedure cannot be precisely determined because C. burnetii's obligate intracellular nature prevents cloning of transformants as colonies on an agar plate. However, in electroporation experiment 5, 13 unique rescue clones were derived. Considering that approximately 1.25 x 10¹⁰ organisms were electroporated, a transformation frequency of approximately 10^–9 can be estimated. This frequency is presumably an underestimate due to the limitations of the rescue cloning procedure and the possible loss over time of transformants harboring deleterious mutations. Earlier C. burnetii transformation experiments utilized resistance to ampicillin as a method of positive selection (36). However, long-term selection resulted in spontaneous mutation to ampicillin resistance by bacteria lacking the introduced β-lactamase gene (36). Here, we show that Cm resistance is a suitable selectable marker due to its ability to prevent C. burnetii growth at a concentration (5 µg/ml) that does not result in obvious host cell toxicity. No growth of Cm-resistant organisms was observed in control electroporation experiments, suggesting that spontaneous mutation to Cm resistance by C. burnetii is rare or does not occur.

In summary, this report details the first successful random transposon mutagenesis of the C. burnetii genome. Saturation level gene inactivation and phenotyping to define the gene complement essential for intracellular replication would be difficult considering our current transformation efficiencies and the time-consuming nature of generating C. burnetii clones. However, one immediate application of the Himar1 system is introduction into the C. burnetii genome of an inducible or constitutively expressed transgene for complementation studies or expression of dominant-negative proteins. The transformation parameters described here will aid development of additional genetic tools, most importantly site-specific allelic exchange. Moreover, mCherry-expressing C. burnetii will have utility in intracellular trafficking studies.

 
We thank John Carlson and Phil Stewart for critical reading of the manuscript and Anita Mora and Gary Hettrick for graphic illustrations.

This research was supported by the Intramural Research Program of the National Institutes of Health, National Institute of Allergy and Infectious Diseases.

 
* Corresponding author. Mailing address: Laboratory of Intracellular Parasites, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, 903 S. 4th St., Hamilton, MT 59840. Phone: (406) 375-9695. Fax: (406) 375-9380. E-mail: rheinzen{at}niaid.nih.gov

Published ahead of print on 29 December 2008.

Supplemental material for this article may be found at http://jb.asm.org/.

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Journal of Bacteriology, March 2009, p. 1369-1381, Vol. 191, No. 5
0021-9193/09/$08.00+0     doi:10.1128/JB.01580-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

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